Cascade Biocatalysis E-Book

• Herausgeber: Wiley-VCH
• Kategorie: Wissenschaft und neue Technologien
• Sprache: Englisch

Seitenzahl: 809

Veröffentlichungsjahr: 2014

Beschreibung

This ready reference presents environmentally friendly and stereoselective methods of modern biocatalysis. The experienced and renowned team of editors have gathered top international authors for this book. They cover such emerging topics as chemoenzymatic methods and multistep enzymatic reactions, while showing how these novel methods and concepts can be used for practical applications. Multidisciplinary topics, including directed evolution, dynamic kinetic resolution, and continuous-flow methodology are also discussed.

From the contents:
* Directed Evolution of Ligninolytic Oxidoreductases: from Functional Expression to Stabilization and Beyond
* New Trends in the In Situ Enzymatic Recycling of NAD(P)(H) Cofactors
* Monooxygenase-Catalyzed Redox Cascade Biotransformations
* Biocatalytic Redox Cascades Involving w-Transaminases
* Multi-Enzyme Systems and Cascade Reactions Involving Cytochrome P450 Monooxygenases
* Chemo-Enzymatic Cascade Reactions for the Synthesis of Glycoconjugates
* Synergies of Chemistry and Biochemistry for the Production of Beta-Amino Acids
* Racemizable Acyl Donors for Enzymatic Dynamic Kinetic Resolution
* Stereoselective Hydrolase-Catalyzed Processes in Continuous-Flow Mode
* Perspectives on Multienzyme Process Technology
* Nitrile Converting Enzymes Involved in Natural and Synthetic Cascade Reactions
* Mining Genomes for Nitrilases
* Key-Study on the Kinetic Aspects of the In-Situ NHase/AMase Cascade System of M. imperiale Resting Cells for Nitrile Bioconversion
* Enzymatic Stereoselective Synthesis of Beta-Amino Acids
* New Applications of Transketolase: Cascade Reactions for Assay Development
* Aldolases as Catalyst for the Synthesis of Carbohydrates and Analogs
* Enzymatic Generation of Sialoconjugate Diversity
* Methyltransferases in Biocatalysis
* Chemoenzymatic Multistep One-Pot Processes

Leseprobe

Table of Contents

Cover

Related Titles

Title Page

Copyright

List of Contributors

Preface

Chapter 1: Directed Evolution of Ligninolytic Oxidoreductases: from Functional Expression to Stabilization and Beyond

1.1 Introduction

1.2 Directed Molecular Evolution

1.3 The Ligninolytic Enzymatic Consortium

1.4 Directed Evolution of Laccases

1.5 Directed Evolution of Peroxidases and Peroxygenases

1.6

Saccharomyces cerevisiae

Biomolecular Tool Box

1.7 Conclusions and Outlook

Acknowledgments

Abbreviations

References

Chapter 2: New Trends in the In Situ Enzymatic Recycling of NAD(P)(H) Cofactors

2.1 Introduction

2.2 Recent Advancements in the Enzymatic Methods for the Recycling of NAD(P)(H) Coenzymes and Novel Regeneration Systems

2.3 Conclusions

Acknowledgments

References

Chapter 3: Monooxygenase-Catalyzed Redox Cascade Biotransformations

3.1 Introduction

References

Chapter 4: Biocatalytic Redox Cascades Involving ω-Transaminases

4.1 Introduction

4.2 General Features of ω-Transaminases

4.3 Linear Cascade Reactions Involving ω-Transaminases

4.4 Concluding Remarks

References

Chapter 5: Multi-Enzyme Systems and Cascade Reactions Involving Cytochrome P450 Monooxygenases

5.1 Introduction

5.2 Physiological Cascade Reactions Involving P450s

5.3 Artificial Cascade Reactions Involving P450s

5.4 Conclusions and Outlook

References

Chapter 6: Chemo-Enzymatic Cascade Reactions for the Synthesis of Glycoconjugates

6.1 Introduction

6.2 Sequential Syntheses

6.3 One-Pot Syntheses

6.4 Convergent Syntheses

6.5 Conclusion

Acknowledgment

References

Chapter 7: Synergies of Chemistry and Biochemistry for the Production of β-Amino Acids

7.1 Introduction

7.2 Dihydropyrimidinase

7.3

N

-Carbamoyl-β-Alanine Amidohydrolase

7.4 Bienzymatic System for β-Amino Acid Production

7.5 Conclusions and Outlook

Acknowledgments

References

Chapter 8: Racemizable Acyl Donors for Enzymatic Dynamic Kinetic Resolution

8.1 Introduction

8.2 The Tools

8.3 Applications of DKR to Acyl Compounds

8.4 Conclusions

Acknowledgments

References

Chapter 9: Stereoselective Hydrolase-Catalyzed Processes in Continuous-Flow Mode

9.1 Introduction

9.2 Enzyme-Catalyzed Stereoselective Reactions in Continuous-Flow Systems

9.3 Outlook and Perspectives

References

Chapter 10: Perspectives on Multienzyme Process Technology

10.1 Introduction

10.2 Multienzyme System Classification

10.3 Biocatalyst Options

10.4 Reactor Options

10.5 Process Development

10.6 Process Modeling

10.7 Future

10.8 Concluding Remarks

References

Chapter 11: Nitrile Converting Enzymes Involved in Natural and Synthetic Cascade Reactions

11.1 Introduction

11.2 Natural Cascades

11.3 Artificial Cascades

11.4 Conclusions and Future Use of These Enzymes

Acknowledgments

References

Chapter 12: Mining Genomes for Nitrilases

12.1 Strategies in Nitrilase Search

12.2 Diversity of Nitrilase Sequences

12.3 Structure–Function Relationships

12.4 Enzyme Properties and Applications

12.5 Conclusions

Acknowledgment

References

Chapter 13: Key-Study on the Kinetic Aspects of the In Situ NHase/AMase Cascade System of M. imperiale Resting Cells for Nitrile Bioconversion

13.1 Introduction

13.2 The Temperature Effect on the NHase–Amidase Bi-Enzymatic Cascade System

13.3 Effect of Nitrile Concentration on NHase Activity and Stability

13.4 Effect of Nitrile on the AMase Activity and Stability

13.5 Concluding Remarks

Acknowledgments

References

Chapter 14: Enzymatic Stereoselective Synthesis of β-Amino Acids

14.1 Introduction

14.2 Preparation of β-Amino Acids

14.3 Nitrile Hydrolysis Enzymes

14.4 Conclusion

Acknowledgments

References

Chapter 15: New Applications of Transketolase: Cascade Reactions for Assay Development

15.1 Introduction

15.2 Cascade Reactions for Assaying Transketolase Activity

In Vitro

15.3 Cascade Reactions for Assaying Transketolase Activity by

In Vivo

Selection

15.4 Conclusion

References

Chapter 16: Aldolases as Catalyst for the Synthesis of Carbohydrates and Analogs

16.1 Introduction

16.2 Iminocyclitol and Aminocyclitol Synthesis

16.3 Carbohydrates and Other Polyhydroxylated Compounds

16.4 Conclusions

Acknowledgments

References

Chapter 17: Enzymatic Generation of Sialoconjugate Diversity

17.1 Introduction

17.2 A Generic Strategy for the Synthesis of Sialoconjugate Libraries

17.3 Cascade Synthesis of neo-Sialoconjugates

17.4 Conclusions

Acknowledgments

References

Chapter 18: Methyltransferases in Biocatalysis

18.1 Introduction

18.2 SAM-Dependent Methyltransferases

18.3 Conclusion and Outlook

Abbreviations

Acknowledgement

References

Chapter 19: Chemoenzymatic Multistep One-Pot Processes

19.1 Introduction: Why Chemoenzymatic Cascades and Why One-Pot Processes?

19.2 Concepts of Chemoenzymatic Processes

19.3 Combination of Substrate Isomerization and their Derivatization with Chemo- and Biocatalysts Resulting in Dynamic Kinetic Resolutions and Related Processes

19.4 Combination of Substrate Synthesis (Without Isomerization) and Derivatization Step(s)

19.5 Conclusion and Outlook

References

Index

End User License Agreement

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Guide

Cover

Table of Contents

Preface

Chapter 1: Directed Evolution of Ligninolytic Oxidoreductases: from Functional Expression to Stabilization and Beyond

List of Illustrations

Figure 1.1

Figure 1.2

Figure 1.3

Figure 1.4

Figure 1.5

Scheme 2.1

Scheme 2.2

Scheme 2.3

Scheme 2.4

Figure 2.1

Scheme 2.5

Scheme 2.6

Scheme 3.1

Scheme 3.2

Scheme 3.3

Scheme 3.4

Scheme 3.5

Scheme 3.6

Scheme 3.7

Scheme 3.8

Scheme 3.9

Scheme 3.10

Scheme 3.11

Scheme 3.12

Scheme 3.13

Scheme 3.14

Scheme 3.15

Scheme 3.16

Scheme 3.17

Scheme 3.18

Scheme 3.19

Scheme 3.20

Scheme 4.1

Scheme 4.2

Scheme 4.3

Scheme 4.4

Scheme 4.5

Scheme 4.6

Scheme 4.7

Scheme 4.8

Scheme 4.9

Scheme 4.10

Scheme 4.11

Scheme 4.12

Scheme 4.13

Scheme 4.14

Scheme 4.15

Scheme 4.16

Scheme 5.1

Scheme 5.2

Figure 5.1

Scheme 5.3

Scheme 5.4

Scheme 5.5

Scheme 5.6

Scheme 5.7

Scheme 5.8

Scheme 5.9

Scheme 5.10

Scheme 5.11

Scheme 5.12

Scheme 5.13

Scheme 5.14

Scheme 5.15

Scheme 5.16

Scheme 5.17

Scheme 5.18

Scheme 5.19

Scheme 5.20

Scheme 5.21

Scheme 5.22

Scheme 5.23

Scheme 5.24

Figure 5.2

Figure 5.3

Scheme 5.25

Scheme 5.26

Scheme 5.27

Scheme 5.28

Scheme 5.29

Scheme 5.30

Scheme 6.1

Scheme 6.2

Scheme 6.3

Figure 6.1

Figure 6.2

Figure 6.3

Figure 6.4

Figure 6.5

Scheme 6.4

Scheme 6.5

Figure 6.6

Figure 6.7

Figure 6.8

Figure 6.9

Figure 7.1

Figure 7.2

Figure 7.3

Figure 7.4

Figure 7.5

Figure 7.6

Scheme 8.1

Scheme 8.2

Scheme 8.3

Figure 8.1

Scheme 8.4

Scheme 8.5

Scheme 8.6

Scheme 8.7

Scheme 8.8

Scheme 8.9

Scheme 8.10

Figure 8.2

Figure 8.3

Figure 9.1

Figure 9.2

Figure 9.3

Figure 9.4

Figure 9.5

Figure 9.6

Figure 9.7

Figure 9.8

Figure 9.9

Figure 9.10

Figure 10.1

Figure 10.2

Figure 10.3

Figure 11.1

Figure 11.2

Figure 11.3

Figure 11.4

Figure 11.5

Figure 11.6

Figure 11.7

Figure 11.8

Figure 11.9

Figure 11.10

Figure 11.11

Figure 11.12

Figure 11.13

Figure 11.14

Figure 11.15

Figure 11.16

Figure 12.1

Figure 12.2

Figure 13.1

Figure 13.2

Figure 13.3

Figure 13.4

Figure 13.5

Figure 13.6

Figure 14.1

Figure 14.2

Scheme 14.1

Scheme 14.2

Scheme 14.3

Scheme 14.4

Scheme 14.5

Scheme 15.1

Scheme 15.2

Scheme 15.3

Scheme 15.4

Scheme 15.5

Scheme 15.6

Scheme 15.7

Scheme 15.8

Scheme 15.9

Scheme 15.10

Scheme 15.11

Scheme 15.12

Figure 15.1

Scheme 15.13

Scheme 15.14

Figure 15.15

Figure 15.2

Figure 15.3

Figure 15.4

Scheme 15.16

Scheme 15.17

Scheme 15.18

Figure 15.5

Scheme 15.19

Scheme 15.20

Scheme 16.1

Scheme 16.2

Figure 16.1

Scheme 16.3

Scheme 16.4

Figure 16.2

Scheme 16.5

Scheme 16.6

Scheme 16.7

Scheme 16.8

Scheme 16.9

Figure 17.1

Scheme 17.1

Figure 17.2

Scheme 17.2

Scheme 17.3

Scheme 17.4

Scheme 17.5

Scheme 17.6

Scheme 17.7

Scheme 17.8

Scheme 17.9

Scheme 17.10

Scheme 17.11

Figure 17.3

Figure 17.4

Scheme 17.12

Scheme 17.13

Scheme 17.14

Scheme 17.15

Scheme 17.16

Scheme 17.17

Scheme 17.18

Scheme 17.19

Scheme 17.20

Scheme 17.21

Scheme 17.22

Scheme 17.23

Figure 18.1

Figure 18.2

Scheme 18.1

Scheme 18.2

Scheme 18.3

Scheme 18.4

Scheme 18.5

Scheme 18.6

Scheme 18.7

Scheme 18.8

Figure 18.3

Scheme 18.9

Figure 18.4

Figure 18.5

Figure 18.6

Scheme 19.1

Scheme 19.2

Scheme 19.3

Scheme 19.4

Scheme 19.5

Scheme 19.6

Scheme 19.7

Scheme 19.8

Scheme 19.9

Scheme 19.10

Scheme 19.11

Scheme 19.12

Scheme 19.13

Scheme 19.14

Scheme 19.15

Scheme 19.16

Scheme 19.17

Scheme 19.18

Scheme 19.19

Scheme 19.20

Scheme 19.21

Scheme 19.22

Scheme 19.23

Scheme 19.24

Scheme 19.25

Scheme 19.26

Scheme 19.27

Scheme 19.28

Scheme 19.29

Scheme 19.30

List of Tables

Table 2.1

Table 4.1

Table 4.2

Table 4.3

Table 4.4

Table 4.5

Table 7.1

Table 7.2

Table 8.1

Table 8.2

Table 8.3

Table 8.4

Table 8.5

Table 8.6

Table 8.7

Table 9.1

Table 9.2

Table 9.3

Table 9.4

Table 9.5

Table 9.6

Table 10.1

Table 10.2

Table 10.3

Table 12.1

Table 13.1

Table 13.2

Table 14.1

Table 14.2

Table 14.3

Table 15.1

Table 15.2

Table 15.3

Table 15.4

Table 16.1

Table 17.1

Table 17.2

Table 18.1

Table 18.2

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Edited by Sergio Riva and Wolf-Dieter Fessner

Cascade Biocatalysis

Integrating Stereoselective and Environmentally Friendly Reactions

The Editors

Dr. Sergio Riva

Istituto di Chimica del Riconoscimento Molecolare

CNR

Via Mario Bianco 9

20131 Milano

Italy

Prof. Wolf-Dieter Fessner

TU Darmstadt

Institut f$\ddot{\hbox{u}}$r Organische Chemie und \hbox{Biochemie}

Petersenstr. 22

64287 Darmstadt

Germany

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List of Contributors

Miguel Alcalde

Institute of Catalysis, CSIC

Department of Biocatalysis

C/Marie Curie n°2

Cantoblanco

28049 Madrid

Spain

Antonio O. Ballesteros

Institute of Catalysis, CSIC

Department of Biocatalysis

C/Marie Curie n°2

Cantoblanco

28049 Madrid

Spain

Moira Bode

University of the Witwatersrand

Molecular Sciences Institute School of Chemistry

PO Wits

Johannesburg

South Africa

Zoltán Boros

Budapest University of Technology and Economics

Department for Organic Chemistry and Technology

Szt Gellért tér 4

H-1111 Budapest

Hungary

Dean Brady

University of the Witwatersrand

School of Chemistry, Molecular Sciences Institute

PO Wits

Johannesburg

South Africa

and

CSIR Biosciences

Scientia campus

CSIR Building 18

Meiring Naude Road

Pretoria, 0184

South Africa

Jordi Bujons

Instituto de Química Avanzada de Cataluña IQAC-CSIC

Dept Chemical Biology and Molecular Modeling

Biotransformation and Bioactive Molecules Group

Jordi Girona 18-26

Barcelona

Spain

Susana Camarero

CSIC, Centro de Investigaciones Biológicas

Ramiro de Maeztu 9

E-28040 Madrid

Spain

Laura Cantarella

University of Cassino and of Lazio Meridionale

Department of Civil and Mechanical Engineering

via Di Biasio 43

Cassino (FR)

Italy

Maria Cantarella

University of L'Aquila

Department of Industrial and Information Engineering and Economics

via Giovanni Gronchi n.18-Nucleo industriale di Pile

L'Aquila

Italy

Franck Charmantray

Université Blaise Pascal

Institut de Chimie de Clermont-Ferrand (ICCF)

UMR CNRS 6296

BP 10448, F-63177 Aubière

France

Varsha Chhiba

CSIR Biosciences

Scientia campus

CSIR Building 18

Meiring Naude Road

Pretoria, 0184

South Africa

Pere Clapés

Instituto de Química Avanzada de Cataluña IQAC-CSIC

Dept Chemical Biology and Molecular Modeling

Biotransformation and Bioactive Molecules Group

Jordi Girona 18-26

Barcelona

Spain

Josefa María Clemente-Jiménez

Universidad de Almería

Departamento de Química y Física

Carretera de Sacramento S/N

Edificio C.I.T.E. I

La Cañada de San Urbano

Almería

Spain

Nicola D'Antona

CNR National Research Council of Italy

Institute of Biomolecular Chemistry

Via P. Gaifami 18

Catania

Italy

Martin Dippe

Leibniz Institute of Plant Biochemistry

Weinberg 3

D-06120 Halle

Germany

Lothar Elling

RWTH Aachen University

Department of Biotechnology and Helmholtz-Institute for Biomedical Engineering

Worringer Weg 1

Aachen

Germany

Erica Elisa Ferrandi

Istituto di Chimica del Riconoscimento Molecolare C. N. R.

Via Mario Bianco 9

Milano

Italy

Wolf-Dieter Fessner

Technische Universität Darmstadt

Department of Organic Chemistry and Biochemistry

Petersenstr 22

D-64287 Darmstadt

Germany

Eva Garcia-Ruiz

Institute of Catalysis, CSIC

Department of Biocatalysis

C/Marie Curie n°2

Cantoblanco

28049 Madrid

Spain

David Gonzalez-Perez

Institute of Catalysis, CSIC

Department of Biocatalysis

C/Marie Curie n°2

Cantoblanco

28049 Madrid

Spain

Harald Gröger

Bielefeld University

Faculty of Chemistry

Universitätsstr 25

Bielefeld

Germany

Mandana Gruber-Khadjawi

ACIB GmbH

c/o Graz University of Technology

Institute of Organic Chemistry

Stremayrgasse 9

Graz

Austria

Ning He

Technische Universität Darmstadt

Department of Organic Chemistry and Biochemistry

Petersenstr 22

D-64287 Darmstadt

Germany

Laurence Hecquet

Université Blaise Pascal

Institut de Chimie de Clermont-Ferrand (ICCF)

UMR CNRS 6296

BP 10448, F-63177 Aubière

France

Virgil Hélaine

Université Blaise Pascal

Institut de Chimie de Clermont-Ferrand (ICCF)

UMR CNRS 6296

BP 10448, F-63177 Aubière

France

Gábor Hornyánszky

Budapest University of Technology and Economics

Department for Organic Chemistry and Technology

Szt Gellért tér 4

H-1111 Budapest

Hungary

Werner Hummel

Heinrich-Heine-University of Düsseldorf

Research Centre Jülich

Institute of Molecular Enzyme Technology

Stetternicher Forst

Jülich

Germany

Jesús Joglar

Instituto de Química Avanzada de Cataluña IQAC-CSIC

Dept Chemical Biology and Molecular Modeling

Biotransformation and Bioactive Molecules Group

Jordi Girona 18-26

Barcelona

Spain

Marion Knorst

Technische Universität Darmstadt

Department of Organic Chemistry and Biochemistry

Petersenstr 22

D-64287 Darmstadt

Germany

Wolfgang Kroutil

University of Graz

Institute of Chemistry

Heinrichstr 28

Graz

Austria

Bastian Lange

RWTH Aachen University

Department of Biotechnology and Helmholtz-Institute for Biomedical Engineering

Worringer Weg 1

Aachen

Germany

Francisco Javier Las Heras-Vázquez

Universidad de Almería

Departamento de Química y Física

Carretera de Sacramento S/N

Edificio C.I.T.E. I

La Cañada de San Urbano

Almería

Spain

Angel T. Martínez

CSIC, Centro de Investigaciones Biológicas

Ramiro de Maeztu 9

E-28040 Madrid

Spain

Sergio Martínez-Rodríguez

Universidad de Almería

Departamento de Química y Física

Carretera de Sacramento S/N

Edificio C.I.T.E. I

La Cañada de San Urbano

Almería

Spain

Ludmila Martínková

Academy of Sciences of the Czech Republic

Institute of Microbiology

Laboratory of Biotransformation

Videnská 1083

CZ-142 20 Prague

Czech Republic

Diana M. Mate

Institute of Catalysis, CSIC

Department of Biocatalysis

C/Marie Curie n°2

Cantoblanco

28049 Madrid

Spain

Kgama Mathiba

CSIR Biosciences

Scientia campus

CSIR Building 18

Meiring Naude Road

Pretoria, 0184

South Africa

Marko D. Mihovilovic

Vienna University of Technology

Institute of Applied Synthetic Chemistry

Getreidemarkt 9/163-OC

A-1060 Vienna

Austria

Patricia Molina-Espeja

Institute of Catalysis, CSIC

Department of Biocatalysis

C/Marie Curie n°2

Cantoblanco

28049 Madrid

Spain

Daniela Monti

Istituto di Chimica del Riconoscimento Molecolare C. N. R.

Via Mario Bianco 9

Milano

Italy

József Nagy

Budapest University of Technology and Economics

Department for Organic Chemistry and Technology

Szt Gellért tér 4

H-1111 Budapest

Hungary

Linda G. Otten

Delft University of Technology

Department of Biotechnology

Biocatalysis

Julianalaan 136

BL Delft

The Netherlands

Fabrizia Pasquarelli

University of L'Aquila

Department of Industrial and Information Engineering and Economics

via Giovanni Gronchi n.18-Nucleo industriale di Pile

L'Aquila

Italy

László Poppe

Budapest University of Technology and Economics

Department for Organic Chemistry and Technology

Szt Gellért tér 4

H-1111 Budapest

Hungary

Nina Richter

University of Graz

Institute of Chemistry

Heinrichstr 28

Graz

Austria

Sergio Riva

Istituto di Chimica del Riconoscimento Molecolare

N. R.

Via Mario Bianco 9

Milano

Italy

Felipe Rodríguez-Vico

Universidad de Almería

Departamento de Química y Física

Carretera de Sacramento S/N

Edificio C.I.T.E. I

La Cañada de San Urbano

Almería

Spain

Ruben R. Rosencrantz

RWTH Aachen University

Department of Biotechnology and Helmholtz-Institute for Biomedical Engineering

Worringer Weg 1

Aachen

Germany

Florian Rudroff

Vienna University of Technology

Institute of Applied Synthetic Chemistry

Getreidemarkt 9/163-OC

A-1060 Vienna

Austria

Paloma A. Santacoloma

Technical University of Denmark (DTU)

Department of Chemical and Biochemical Engineering

Søltofts Plads

Kgs. Lyngby

Denmark

Sebastian Schulz

Heinrich-Heine University Düsseldorf

Institute of Biochemistry

Universitätsstr 1

Düsseldorf

Germany

Robert C. Simon

University of Graz

Institute of Chemistry

Heinrichstr 28

Graz

Austria

Agata Spera

University of L'Aquila

Department of Industrial and Information Engineering and Economics

via Giovanni Gronchi n.18-Nucleo industriale di Pile

L'Aquila

Italy

Andreas Stolz

University of Stuttgart

Institute of Microbiology

Allmandring 31

Stuttgart

Germany

Martin Tengg

ACIB GmbH

c/o Graz University of Technology

Institute of Molecular Biotechnology

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Graz

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Davide Tessaro

Politecnico di Milano

Department of Chemistry Materials and Chemical Engineering ``G. Natta"

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Milano

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and

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University Center for Protein Biotechnology

via Mancinelli, 7

Milano

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Peter Unruh

Technische Universität Darmstadt

Department of Organic Chemistry and Biochemistry

Petersenstr 22

D-64287 Darmstadt

Germany

Vlada B. Urlacher

Heinrich-Heine University Düsseldorf

Institute of Biochemistry

Universitätsstr 1

Düsseldorf

Germany

Fred van Rantwijk

Delft University of Technology

Department of Biotechnology

Biocatalysis

Julianalaan 136

BL Delft

The Netherlands

Ludger Wessjohann

Leibniz Institute of Plant Biochemistry

Weinberg 3

D-06120 Halle

Germany

John M. Woodley

Technical University of Denmark (DTU)

Department of Chemical and Biochemical Engineering

Søltofts Plads

Kgs. Lyngby

Denmark

Dong Yi

Technische Universität Darmstadt

Department of Organic Chemistry and Biochemistry

Petersenstr 22

D-64287 Darmstadt

Germany

Preface

Sustainability is one of the key issues to enhance, or at least maintain, the quality of life in our modern society. As it has been codified in 1987 in an official UN document, a “sustainable development is development that meets the needs of the present without compromising the ability of future generations to meet their own needs”. Applied to chemical processes, sustainability has generated the concept of Green Chemistry, for which guidelines have been summarized as the well-known Twelve Principles of Green Chemistry1.

In Europe, this effort has been recognized at the institutional level: the European Technology Platform for Sustainable Chemistry (SusChem, http://www.suschem.org) was created in 2004 with the main objective to revitalize and inspire the European chemistry research, development, and innovation in a sustainable way. Industrial Biotechnology, also known as White Biotechnology, is one of the three pillars that support sustainable chemistry nowadays and that are expected to support it even more profoundly in the future. It is defined as “the use of enzymes and micro-organisms to make efficient and sustainable products in sectors as diverse as chemicals, plastics, food and feed, detergents, paper and pulp, textiles or bioenergy.”

Although long and reiterating, this introduction is meant to raise the awareness that the roots and the branches of biocatalysis – as well as its fruits! – are deeply embedded in modern synthetic chemistry. In fact, the majority of the above-mentioned Principles of Green Chemistry (PGC) fit perfectly with the peculiar properties and synthetic application of enzymes, which are Nature's catalysts. The contributions collected in this book offer a convincing testimony that biocatalysis is highly qualified to contribute to the development of future sustainable technologies. Enzymes are highly efficient catalysts offering superior selectivity (PGC #9), thereby meeting criteria for atom economy by maximizing the incorporation of starting materials into the final product (PGC #2) while avoiding unnecessary and unproductive derivatization, such as the use of temporary protection groups (PGC #8). Such steps are unavoidable when using conventional synthetic chemistry approaches and require additional reagents and generate waste materials, particularly when utilizing multifunctionalized, bio-based renewable feedstocks (PGC #7). Inherently, enzymes are biodegradable (PGC #10) and innocuous to the environment (PGC #3), not the least because they operate in water as a safe solvent (PGC #5) at ambient temperature and pressure, which minimizes energy consumption (PGC #6).

Cascade Biocatalysis is an effort to imitate the style of chemical conversions occurring in living beings, which are totally different from the traditional use of single enzymes by synthetic chemists in the laboratory for catalyzing isolated transformations. Instead, cells apply multistep synthetic strategies, catalyzed by several enzymes acting sequentially along a pathway, in which a product formed in one reaction in situ becomes the substrate of the next catalyst. This is possible because of the very similar mild reaction conditions under which most enzymes operate, which facilitates their combination and allows effective strategies of reaction engineering, for example, to shift unproductive equilibria by coupling to thermodynamically favored processes for overall high conversion and economic efficiency.

This concept has recently been recognized as the major focus for a series of international symposia on Multistep Enzyme-Catalyzed Processes, the last symposium having just been celebrated in Madrid in April 2014. Research in this area has also been coordinated within the activities of the European Union funded COST network CM0701 entitled Cascade Chemoenzymatic Processes – New Synergies Between Chemistry and Biochemistry (2008–2012; http://www.cost-cascat.polimi.it). This handbook brings together contributions from scientists deeply involved in the activities of this COST action as well as complementary chapters on related research from additional authors, who are well known for their seminal work in this contemporary research field. The topics covered in the chapters span from examples related to integrated applications of cofactor-dependent oxidoreductases to the exploitation of transferases; from the multistep modification of the nitrile functional group to the synthesis of complex carbohydrates; and from developments of new dynamic kinetic resolution processes to intricate examples of chemoenzymatic multistep one-pot procedures.

We would like to thank all the authors who, despite their busy schedules, have participated in this project to share their expertise with the future readers of this book. Thanks are also due to Elke Maase and Stefanie Volk at Wiley-VCH Publishers, for their careful editorial support and for their continuous goad in order to meet assigned deadlines.

Finally, we hope that our readers will find this volume useful as a stimulating source of ideas for their own research and/or teaching activities.

Sergio Riva

Wolf-Dieter Fessner

1

P. Anastas, J.C. Warner,

Green Chemistry: Theory and Practice

, Oxford University Press (2008).

Chapter 1Directed Evolution of Ligninolytic Oxidoreductases: from Functional Expression to Stabilization and Beyond

Eva Garcia-Ruiz, Diana M. Mate, David Gonzalez-Perez, Patricia Molina-Espeja, Susana Camarero, Angel T. Martínez, Antonio O. Ballesteros, and Miguel Alcalde

1.1 Introduction

The ligninolytic enzymatic consortium, formed mainly by nonspecific oxidoreductases (laccases, peroxidases, and H2O2-supplying oxidases), is a potentially powerful multipurpose tool for industrial and environmental biotechnology. In nature, these enzymes are typically produced by basidiomycete white-rot fungi that are involved in lignin decay. Thanks to their broad substrate specificity, high redox potential, and minimal requirements, these enzymes have many potential applications in the field of green chemistry, including the production of biofuels, bioremediation, organic syntheses, pulp biobleaching, food and textile industries, and the design of bionanodevices. The implementation of this enzymatic armoury in different biotechnological sectors has been hampered by the lack of appropriate molecular instruments (including heterologous hosts for directed evolution) with which to improve their properties. Over the last 10 years, a wealth of directed evolution strategies in combination with hybrid approaches has emerged in order to adapt these oxidoreductases to the drastic conditions associated with many biotechnological settings (e.g., high temperatures, the presence of organic co-solvents, extreme pHs, the presence of inhibitors). This chapter summarizes all efforts and endeavors to convert these ligninolytic enzymes into useful biocatalysts by means of directed evolution: from functional expression to stabilization and beyond.

1.2 Directed Molecular Evolution

Enzymes are versatile biomolecules that exhibit a large repertory of functions acquired over millions of years of natural evolution. Indeed, they are the fastest known catalysts (accelerating chemical reactions as much as 1019-fold) and are environmentally friendly molecules, working efficiently at mild temperatures, in water, and releasing few by-products. Moreover, they can exhibit high enantioselectivity and chemoselectivity. Nonetheless, when an enzyme is removed from its natural environment and introduced into a specific biotechnological location (e.g., the transformation of a hydrophobic compound in the presence of co-solvents or at high temperatures), its molecular structure may not tolerate the extreme operational conditions and may unfold becoming inactive. Unfortunately, the enzymes that cells use to regulate strict metabolic pathways and that promote fitness and survival in nature are not always applicable to the harsh requirements of many industrial processes.

The development of the polymerase chain reaction (PCR) in the early 1980s heralded a biotechnological revolution for protein engineers, allowing us for the first time to manipulate and design enzymes by site-directed mutagenesis supported by known protein structures: the so-called rational design. However, further advances were frustrated owing to the limited understanding of protein function and the lack of protein structures available at the time. Nevertheless, the following decade saw a second biotechnological revolution with the development of directed molecular evolution. This powerful protein engineering tool does not require prior knowledge of protein structure to enhance the known features or to generate novel enzymatic functions, which are not generally required in natural environments. The key events of natural evolution (random mutation, DNA recombination, and selection) are recreated in the laboratory, permitting scientifically interesting and technologically useful enzymes to be designed [1–3]. Diversity is generated by introducing random mutations and/or recombination in the gene encoding a specific protein [4, 5]. In this process, the best performers in each round of evolution are selected and used as the parental types in a new round, a cycle that can be repeated as many times as necessary until a biocatalyst that exhibits the desired traits is obtained: for example, improved stability at high temperatures, extreme pHs, or in the presence of nonconventional media such as organic solvents or ionic fluids; novel catalytic activities; improved specificities and/or modified enantioselectivities; and heterologous functional expression [6–8] (Figure 1.1). Of great interest is the use of directed evolution strategies to engineer ligninolytic oxidoreductases while employing rational approaches to understand the mechanisms underlying each newly evolved property.

Figure 1.1 Directed molecular evolution. The basic premises to carry out a successful directed evolution experiment are (i) a robust heterologous expression system (typically S. cerevisiae or E. coli); (ii) a reliable high-throughput (HT)-screening assay; and (iii) the use of different molecular tools for the generation of DNA diversity.

1.3 The Ligninolytic Enzymatic Consortium

Lignin is the most abundant natural aromatic polymer and the second most abundant component of plant biomass after cellulose. As a structural part of the plant cell wall, lignin forms a complex matrix that protects cellulose and hemicellulose chains from microbial attack and hence from enzymatic hydrolysis. This recalcitrant and highly heterogeneous biopolymer is synthesized by the dehydrogenative polymerization of three precursors belonging to the p-hydroxycinnamyl alcohol group: p-coumaryl, coniferyl, and sinapyl alcohols [9]. As one-third of the carbon fixed as lignocellulose is lignin, its degradation is considered a key step in the recycling of carbon in the biosphere and in the use of the plant biomass for biotechnological purposes [10, 11]. Lignin is modified and degraded to different extents by a limited number of microorganisms, mainly filamentous fungi and bacteria. Lignin degradation by bacteria is somewhat limited and much slower than that mediated by filamentous fungi [12, 13]. Accordingly, the only organisms capable of completing the mineralization of lignin are the white-rot fungi, which produce a white-colored material upon delignification because of the enrichment in cellulose [14, 15].

Through fungal genome reconstructions, recent studies have linked the formation of coal deposits during the Permo-Carboniferous period (∼260 million years ago) with the nascent and evolution of white-rot fungi and their lignin-degrading enzymes [16]. Lignin combustion by white-rot fungi involves a very complex extracellular oxidative system that includes high-redox potential laccases (HRPLs), peroxidases and unspecific peroxygenases (UPOs), H2O2-supplying oxidases and auxiliary enzymes, as well as radicals of aromatic compounds and oxidized metal ions that act as both diffusible oxidants and electron carriers [12, 13, 15, 17]. Although the role of each component of the consortium has been studied extensively, many factors remain to be elucidated (Figure 1.2).

Figure 1.2 General view of the plant cell wall and the action of the ligninolytic enzymatic consortium. The lignin polymer is oxidized by white-rot fungi laccases and peroxidases, producing nonphenolic aromatic radicals (1) and phenoxy radicals (2). Nonphenolic aromatic radicals can suffer nonenzymatic modifications such as aromatic ring cleavage (3), ether breakdown (4), Cα–Cβ cleavage (5), and demethoxylation (6). The phenoxy radicals (2) can repolymerize on the lignin polymer (7) or be reduced to phenolic compounds by AAO (8) (concomitantly with aryl alcohol oxidation). These phenolic compounds can be re-oxidized by fungal enzymes (9). In addition, phenoxy radicals can undergo Cα–Cβ cleavage to produce p-quinones (10). Quinones promote the production of superoxide radicals via redox cycling reactions involving QR, laccases, and peroxidases (11, 12). The aromatic aldehydes released from Cα–Cβ cleavage, or synthesized by fungi, are involved in the production of H2O2 via another redox cycling reaction involving AAD and AAO (13, 14). Methanol resulting from demethoxylation of aromatic radicals (6) is oxidized by MOX to produce formaldehyde and H2O2 (15). Fungi also synthesize glyoxal, which is oxidized by GLX to produce H2O2 and oxalate (16), which in turn chelate Mn3+ ions produced by MnP (17). The Mn3+ chelated with organic acids acts as a diffusible oxidant for the oxidation of phenolic compounds (2). The reduction of ferric ions present in wood is mediated by the superoxide radical (18) and they are re-oxidized by the Fenton reaction (19) to produce hydroxyl radicals, which are very strong oxidizers that can attack the lignin polymer (20). AAO, aryl-alcohol oxidase; AAD, aryl-alcohol dehydrogenase; GLX, glyoxal oxidase; LiP, lignin peroxidase; MnP, manganese peroxidase; MOX, methanol oxidase; QR, quinone reductase; VP, versatile peroxidase.

(Figure adapted from [18, 19].) (Source: Bidlack, J.M. et al. 1992 [18], Fig. 1, p. 1. Reproduced with permission of the Oklahoma Academy of Science.)

Laccases typically oxidize the phenolic units of lignin. Lignin peroxidases (LiPs) oxidize both nonphenolic lignin structures and veratryl alcohol (VA), a metabolite synthesized by fungi that helps LiP to avoid inactivation by H2O2 and whose radical cation may act as a redox mediator [20]. Manganese peroxidases (MnPs) generate Mn3+, which upon chelation with organic acids (e.g., oxalate synthesized by fungi) attacks phenolic lignin structures; in addition, MnP can also oxidize nonphenolic compounds via lipid peroxidation [21]. Versatile peroxidases (VPs) combine the catalytic activities of LiP, MnP, and generic peroxidases to oxidize phenolic and nonphenolic lignin units [22]. Some fungal oxidases produce the H2O2 necessary for the activity of peroxidases. Among them, aryl-alcohol oxidase (AAO) transforms benzyl alcohols to the corresponding aldehydes; glyoxal oxidase (GLX) oxidizes glyoxal producing oxalate, which in turn chelates Mn3+; and then methanol oxidase (MOX) converts methanol into formaldehyde; all the above oxidations are coupled with O2 reduction of H2O2. Other enzymes such as cellobiose dehydrogenase (CDH) have been indirectly implicated in lignin degradation. This is because of CDH ability to reduce both ferric iron and O2-generating hydroxyl radicals via Fenton reaction. These radicals are strong oxidizers that act as redox mediators playing a fundamental role during the initial stages of lignin polymer decay, when the small pore size of the plant cell wall prevents the access of fungal enzymes [23]. The same is true for laccases, whose substrate spectrum can be broadened in the presence of natural mediators to act on nonphenolic parts of lignin [24].

High-redox potential laccases and peroxidases/peroxygenases are of great biotechnological interest [25, 26]. With minimal requirements and high redox potentials (up to +790 mV for laccases and over +1000 mV for peroxidases), these enzymes can oxidize a wide range of substrates, finding potential applications in a variety of areas, which are as follows:

The use of lignocellulosic materials (e.g., agricultural wastes) in the production of second-generation biofuels (bioethanol, biobutanol) or the manufacture of new cellulose-derived and lignin-derived value-added products.

The organic synthesis of drugs and antibiotics, cosmetics and complex polymers, and building blocks.

In nanobiotechnology as (i) biosensors (for phenols, oxygen, hydroperoxides, azides, morphine, codeine, catecholamines, or flavonoids) for clinical and environmental applications; and (ii) biofuel cells for biomedical applications.

In bioremediation: oxidation of polycyclic aromatic hydrocarbons (PAHs), dioxins, halogenated compounds, phenolic compounds, benzene derivatives, nitroaromatic compounds, and synthetic organic dyes.

The food industry: drink processing and bakery products.

The paper industry: pulp biobleaching, pitch control, manufacture of mechanical pulps with low energy cost, and effluent treatment.

The textile industry: remediation of dyes in effluents, textile bleaching (e.g., jeans), modification of dyes and fabrics, detergents.

A few years ago, the engineering and improvement of ligninolytic oxidoreductases was significantly hampered by the lack of suitable heterologous hosts to carry out directed evolution studies. Fortunately, things have changed and several reliable platforms for the directed evolution of ligninolytic peroxidases, peroxygenases, and several medium-redox potential laccases and high-redox potential laccases (HRPLs) have been developed using the budding yeast Saccharomyces cerevisiae. These advances have allowed us, for the first time, to specifically tailor ligninolytic oxidoreductases to address new challenges.

1.4 Directed Evolution of Laccases

Laccases (EC 1.10.3.2) are extracellular glycoproteins that belong to the blue multicopper oxidase family (along with ascorbate oxidase, ceruloplasmin, nitrite reductase, bilirubin oxidase, and ferroxidase). Widely distributed in nature, they are present in plants, fungi, bacteria, and insects [27, 28]. Laccases are green catalysts, which are capable of oxidizing dozens of compounds using O2 from air and releasing H2O as their sole by-product [29–31]. These enzymes harbor one type I copper (T1), at which the oxidation of the substrates takes place, and a trinuclear copper cluster (T2/T3) formed by three additional coppers, one T2 and two T3s, at which O2 is reduced to H2O. The reaction mechanism resembles a battery, storing electrons from the four monovalent oxidation reactions of the reducing substrate required to reduce one molecule of oxygen to two molecules of H2O. Laccases catalyze the transformation of a wide variety of aromatic compounds, including ortho- and para-diphenols, methoxy-substituted phenols, aromatic amines, benzenothiols, and hydroxyindols. Inorganic/organic metal compounds are also substrates of laccases, and it has been reported that Mn2+ is oxidized by laccase to form Mn3+, and organometallic compounds such [Fe(CN)6]2− are also accepted by the enzyme [32]. The range of reducing substrates can be further expanded to nonphenolic aromatic compounds, otherwise difficult to oxidize, by including redox mediators from natural or synthetic sources. Upon oxidation by the enzyme, such mediators act as diffusible electron carriers in the so-called laccase-mediator systems [24].

Later we summarize the main advances made in the directed evolution of this interesting group of oxidoreductases, paying particular attention to fungal laccases.

1.4.1 Directed Evolution of Low-Redox Potential Laccases

Several directed evolution studies of bacterial laccase CotA have successfully improved its substrate specificity and functional expression, modifying its specificities by screening mutant libraries through surface display [33–37]. The advantages of some bacterial laccases include high thermostability and activity at neutral/alkaline pH, although a low-redox potential at the T1 site often precludes their use in certain sectors.

1.4.2 Directed Evolution of Medium-Redox Potential Laccases

The first successful example of the directed evolution of fungal laccase involved the laccase from the thermophile ascomycete Myceliophthora thermophila laccase (MtL). This study led to subsequent directed evolution experiments in S. cerevisiae with several high-redox potential ligninolytic oxidoreductases (see below). MtL was subjected to 10 cycles of directed evolution to enhance its functional expression in S. cerevisiae [38]. The best performing variant of this process (the T2 mutant that harbored 14 mutations) exhibited a total improvement of 170-fold in activity: its expression levels were enhanced 8-fold and the kcat/Km around 22-fold. The H(c2)R mutation at the C-terminal tail of MtL introduced a recognition site for the KEX2 protease of the Golgi compartment, which facilitated its appropriate maturation and secretion by yeast. Using this laccase expression system as a departure point, five further cycles of evolution were performed to make the laccase both active and stable in the presence of organic co-solvents, a property that makes it suitable for many potential applications in organic syntheses and bioremediation [39–42]. The stability variant (the R2 mutant) functioned in high concentrations of co-solvents of different chemical natures and polarities (a promiscuity toward co-solvents that was promoted during the directed evolution [40]). Most of the mutations introduced in the evolutionary process were located at the surface of the protein, establishing new interactions with surrounding residues, which resulted in structural reinforcement. In the course of these 15 generations of evolution for functional expression in yeast [38] and stabilization in the presence of organic co-solvents [40], the laccase shifted its optimum pH toward less acidic values. Fungal laccases that are active at neutral/alkaline pHs are highly desirable for many applications, such as detoxification, pulp biobleaching, biomedical uses, and enzymatic co-factor regeneration. Recently, the MtL-R2 mutant was converted into an alkalophilic fungal laccase [43]. Accordingly, a high-throughput screening (HTS) assay based on the activity ratio at pH 8.0 to 5.0 was used as the main discriminatory factor. Screening the laccase mutant libraries at alkaline pH while conserving activity at acidic pHs led to a shift in the pH activity profiles that was accompanied by improved catalytic efficiency at both pH values (31-fold at pH 7.0 and 12-fold at pH 4.0). The final variant obtained in this evolution experiment (the IG-88 mutant) retained 90% of its activity at pH 4.0–6.0 and 50% at pH 7.0, and some activity was even detected at pH 8.0.

After 20 generations, the successful in vitro evolution of MtL can be attributed to the plasticity and robustness of this thermostable protein, highlighting that there may be an additional margin for further engineering.

1.4.3 Directed Evolution of Ligninolytic High-Redox Potential Laccases (HRPLs)

Two HRPLs from the basidiomycete PM1 laccase (PM1L) and Pycnoporus cinnabarinus laccase (PcL) were subjected to parallel comprehensive directed evolution in order to achieve functional expression in S. cerevisiae while conserving their thermostability [44]. PM1L was tailored during eight cycles of directed evolution combined with semirational/hybrid approaches [45]. The native laccase signal sequence was replaced by the α-factor prepro-leader from S. cerevisiae and it was evolved in conjunction with the mature protein, adjusting both elements for a successful exportation by yeast. After screening over 50 000 clones, this approach led to the generation of a highly active, soluble, and thermostable HRPL. The total improvement in activity achieved was as high as 34 000-fold relative to the parental type, an effect brought about by the synergies established between the evolved prepro-leader and the mature laccase. Several strategies were employed to maintain the stability of the laccase while enhancing its activity and secretion during evolution: (i) screening for stabilizing mutations [46]; (ii) mutational exchange with beneficial PcL mutations; and (iii) mutational recovery of beneficial mutations with a low likelihood of recombination [44, 47].

Figure 1.3 General structure and details of the blood-tolerant laccase (ChU-B mutant). The F396I and F454E mutations are located 7.6 Å away from the T1 Cu site (in the second coordination sphere). The 3D structure model is based on the crystal structure of the Trametes trogii laccase (97% identity, PDB: 2HRG).

The final mutant generated in this process (the OB-1 variant with 15 mutations accumulated both in the prepro-leader and in the mature protein) exhibited secretion levels of ∼8 mg l−1, and it was very active and stable over a range of temperatures (T50 ∼ 73 °C) and pH values, as well as in the presence of organic co-solvents [45]. OB-1 was recently subjected to four further rounds of directed evolution and saturation mutagenesis in order to achieve activity in human blood, a milestone that will allow it to be used in a wide array of exciting biomedical and bioanalytical applications [48]. The inherent inhibition of laccase by the combined action of high NaCl concentrations (∼140 mM) and the alkaline pH (∼7.4) of blood was overcome by using an ad hoc HTS assay in a buffer that simulated blood but lacked coagulating agents and red blood cells. Bearing in mind that HRPLs are not active at neutral pH, the selective pressure was enhanced in successive rounds of evolution, starting at pH 6.5 and finishing at physiological pH. The final laccase mutant obtained (the ChU-B variant) was comprehensively characterized and tested in real human blood samples, revealing the mechanisms underlying this unprecedented improvement. The ChU-B variant conserved a high-redox potential at the T1 site and exhibited the highest tolerance to halides reported for any HRPL (with an increase in the I50 for Cl− from 176 mM to over 1 M with -2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid (ABTS) as the substrate), and it displayed significant activity at neutral pH (retaining 50% and 20% of its activity for 2,6-dimethoxyphenol (DMP) and ABTS, respectively). This was the first successful example of the use of laboratory evolution to optimize an oxidoreductase for enhanced catalysis in blood for biomedical purposes. From a more general point of view, this development is of considerable importance for a wide range of biotechnological sectors (e.g., bioremediation, pulp-kraft biobleaching), and especially in biocatalysis to develop novel green syntheses. With respect to the parental type, the ChU-B variant presented only two extra mutations in the mature protein (F396I and F454E), which were responsible for its activity in blood but compromised its stability (a 10 °C decrease in the T50, Figure 1.3). By individually analyzing F454E and F396I mutations, a shift in the pH profile from 4.0 to 5.0 (with DMP as substrate) was detected along with a considerable increases in the I50 for halides and decreases in T50 values (4.8 and 3.6 °C for both mutations, respectively). When a crossroad is reached between activity and stability, it is difficult to further evolve the protein as it does not tolerate the introduction of new sets of beneficial but destabilizing mutations without compromising its structure and function. We are currently attempting to improve the stability of this variant by introducing new stabilizing mutations, such as A361T and S482L from the 16B10 mutant of PM1L [46]. These results reflect the subtle equilibrium between activity and stability when evolving enzymes in the laboratory for nonnatural functions, consistent with the observations in earlier directed evolution studies. For example, a decrease by 10 °C in the T50 was obtained following the directed evolution of P450 BM-3 from Bacillus megaterium to convert it into an alkane monooxygenase [49–51].

To conclude this mutational pathway, PM1L was sculpted by 12 rounds of directed evolution, in which it accumulated 22 mutations (8 silent) throughout the entire fusion gene. Beneficial mutations that enhanced secretion or activity were located in the signal prepro-leader (5 mutations) and the mature protein (7 mutations), respectively. Significantly, only two mutations located in the second coordination sphere of the T1 copper site conferred tolerance to blood. Therefore, the re-specialization required to adapt the PM1L to such inclement conditions affected only 0.4% of the amino acid sequence.

The evolution of the HRPL from PcL was tackled using a similar approach to that described for PM1L (i.e., joint evolution of the α-factor prepro-leader and the mature protein). Six cycles of directed evolution were performed to obtain an enzyme that could be readily expressed by yeast (with secretion levels of ∼2 mg l−1 [52]). A multiple HTS assay based on the oxidation of natural and synthetic redox mediators was employed to discriminate between mutants with improved activities against phenolic and nonphenolic compounds. The final variant of this process (the 3PO mutant, containing 14 mutations) retained its thermostability while significantly broadening its pH activity profile. Notably, the breakdown in secretion and activity was accomplished by fusing the evolved prepro-leader to the native PcL. The evolved signal sequence improved secretion 40-fold, while the mutations that accumulated in the evolved mature protein were responsible for a ∼14-fold enhancement in the kcat, together with an improved secretion/folding of the enzyme (∼14-fold improvement). The directed evolution of signal peptides to enhance secretion and their additional attachment to nonevolved proteins is a valuable strategy for the directed evolution of other ligninolytic oxidoreductases (unspecific peroxygenases, see below [53]).

The sequence identity between PcL and PM1L is over 77%, which facilitated mutational exchange between the two parallel evolution pathways and allowed us to switch protein sequence blocks to create chimeric proteins of HRPLs with hybrid or even enhanced features. To favor multiple crossover events between laccase scaffolds, in vitro and in vivo DNA recombination methods were combined in a single evolutionary step (see Section 1.6). Chimeras with up to six crossover events per sequence were identified, which generated active laccase hybrids with combined characteristics in terms of substrate affinity, pH activity, and thermostability [54]. Interestingly, some chimeras showed higher thermostabilities than the original laccases, demonstrating the importance of accumulating neutral mutations to create an artificial genetic drift that is beneficial to stabilize the protein structure. Other laccase chimeragenesis experiments have been performed using laccase isoenzymes from Trametes sp. C30, but employing a low-redox potential laccase backbone to construct the chimeric libraries [55].

PcL and PM1L evolution aside, the lcc1 gene from Trametes versicolor laccase (TvL) was evolved in the yeast Yarrowia lipolytica, demonstrating the potential for directed evolution in this host [56]. More recently, the lcc2 gene from TvL expressed by S. cerevisiae was subjected to two rounds of random mutagenesis for improved ionic liquid resistance [57]. In addition, directed evolution experiments have been carried out with HRPLs from Pleurotus ostreatus to enhance the laccase activity in combination with computational approaches [58–60], and with HRPLs from Rigidoporus lignosus to increase functional expression in Pichia pastoris [61]. Recently, the evolved PM1L was analyzed using a computational algorithm to elucidate the physical forces that govern the thermostability of the variant [62]. Indeed, the combination of in silico computational methods (based on Monte Carlo simulations and molecular dynamics) and directed evolution may offer new directions to study evolved enzymes.

1.5 Directed Evolution of Peroxidases and Peroxygenases

Ligninolytic peroxidases (EC 1.11.1) are high-redox potential oxidoreductases belonging to Class II of the plant-fungal-prokaryotic peroxidase superfamily, and they correspond to fungal secreted heme-containing peroxidases. These enzymes contain ∼300 amino acids distributed in 10–12 α-helix and 4–5 short β-structures that are located in two domains. The heme-prosthetic group contains an Fe3+ in the resting state, and the overall structure is supported by four or five disulfide bridges and two structural Ca2+ ions that confer stability to the protein. The general catalytic cycle of ligninolytic peroxidases begins with the oxidation of the enzyme by one molecule of H2O2. This activates the enzyme to Compound I (a two-electron-deficient intermediate), which under turnover conditions is reduced back to the resting state via two successive one-electron oxidation steps. Ligninolytic peroxidases are divided into three types [12, 13, 15, 26, 63, 64]:

Lignin peroxidases (LiP, EC 1.11.1.13) are capable of directly oxidizing model lignin dimers and nonphenolic aromatic compounds, as well as other high-redox potential substrates (including dyes) using VA as redox mediator, through a catalytic tryptophan located at the surface of the protein.

Mn peroxidases (MnP, EC 1.11.1.14) oxidize Mn

2+

to form Mn

3+

, which upon chelation with organic acids can act as a diffusible oxidant for the oxidation of phenolic compounds.

Versatile peroxidases (VP, EC 1.11.1.16) combine the catalytic properties of LiP and MnP, and they exhibit great versatility and biotechnological potential. VP oxidizes typical LiP substrates (e.g., VA, methoxybenzenes, and nonphenolic model lignin compounds), as well as Mn

2+

(the classical MnP substrate). VP contains a manganese binding site similar to that of MnP, and a surface catalytic Trp similar to that of LiP that is involved in the oxidation of high- and medium-redox potential compounds but that also oxidizes azo-dyes and other nonphenolic compounds with high-redox potential in the absence of mediators. VP also contains a third catalytic site, located at the entrance to the heme channel, involved in the oxidation of low- to medium-redox potential compounds (similar to generic (low-redox potential) peroxidases).

As described earlier for ligninolytic laccases, the directed evolution of high-redox potential peroxidases has also been hindered by the absence of suitable heterologous expression systems. Most attempts at directed peroxidase evolution have to date been carried out using generic peroxidases. The Coprinopsis cinerea peroxidase (CIP) was evolved to enhance its operational stability versus temperature, H2O2, and alkaline pH. S. cerevisiae was used as the expression system in the evolution process, and the mutated variants were subsequently overexpressed in Aspergillus oryzae [65]. These in vitro evolution studies were complemented by the resolution of the crystal structures of both wild type and evolved CIP [66]. A few years later, the evolution of horseradish peroxidase (HRP) for functional expression in S. cerevisiae and overexpression in P. pastoris was described, using this system to improve the thermal stability and resistance to H2O2 [67]. A recent report described the evolution of de novo designed proteins with peroxidase activity [68]. With regard to ligninolytic peroxidases, using an in vitro expression system based on Escherichia coli, preliminary attempts were made to enhance the oxidative stability of MnP [69]. Some years later, LiP was evolved to enhance its catalytic rate and stability by both yeast surface display and secretion to the extracellular medium [70, 71].

Figure 1.4 (a) General overview of protein processing, maturation and exocytosis in yeast. (b,c) Processing of the α-factor prepro-leader with/without N-terminal extension (EAEA).

Figure 1.5 Different genetic methods for library creation in S. cerevisiae: (a) IVOE; (b) IvAM; (c) StEP + in vivo DNA shuffling; (d) CLERY; (e) MORPHING; and (f) DNA assembler.

VP was recently evolved for secretion, thermostabilization, and H2O2 resistance ([72, 73] and Gonzalez-Perez, D., et al., unpublished material). First, a fusion gene formed by the α-factor prepro-leader and the mature VP from Pleurotus eryngii was subjected to four cycles of directed evolution to favor functional expression in S. cerevisiae, achieving secretion levels of ∼22 mg l−1. The secretion mutant (R4 variant) harbored four mutations in the mature protein and increased its total VP activity 129-fold relative to the parental type, together with a marked improvement in catalytic efficiency at the heme channel. Although the catalytic Trp was unaltered after evolution, the Mn2+ site was negatively affected by the mutations. Notably, signal leader processing by the STE13 protease at the Golgi compartment was altered as a consequence of the levels of VP expression, retaining the additional N-terminal sequence EAEA (Glu-Ala-Glu-Ala, Figure 1.4). A similar effect was detected with the evolved prepro-leader of the laccase OB-1 [74]. With both enzymes, the engineered N-terminal truncated variants displayed similar biochemical properties to those of their nontruncated counterparts, although their secretion levels were negatively affected, probably owing to the modifications in the acidic environment close to the KEX2 cleavage site. The R4 secretion mutant was used as the departure point to improve thermostability [46, 72] and three additional cycles of evolution led to a more thermostable variant (2-1B), harboring three additional stabilizing mutations. The 2-1B mutant exhibited a T50